REVIEW ARTICLE
Human HAD phosphatases: structure, mechanism, and
roles in health and disease
Annegrit Seifried1, Jörg Schultz2 and Antje Gohla1,3
1 Rudolf Virchow Center for Experimental Biomedicine, University of Würzburg, Germany
2 Biocenter, Department for Bioinformatics, University of Würzburg, Germany
3 Institute for Pharmacology and Toxicology, University of Würzburg, Germany
Keywords
aspartyl transferase; cancer; cardiovascular
disease; HAD phosphatase; metabolism;
neurological disease; Rossmann fold
Correspondence
A. Gohla, Institute for Pharmacology and
Toxicology and Rudolf Virchow Center,
DFG Research Center for Experimental
Biomedicine, University of Würzburg,
Versbacher Str. 9, 97078 Würzburg,
Germany
Fax: +49 931 201 48539
Tel: +49 931 201 48977
E-mail: antje.gohla@virchow.
uni-wuerzburg.de
(Received 29 February 2012, revised 25
April 2012, accepted 14 May 2012)
Phosphatases of the haloacid dehalogenase (HAD) superfamily of hydrolases are an ancient and very large class of enzymes that have evolved to
dephosphorylate a wide range of low- and high molecular weight substrates
with often exquisite specificities. HAD phosphatases constitute approximately one-fifth of all human phosphatase catalytic subunits. While the
overall sequence similarity between HAD phosphatases is generally very
low, family members can be identified based on the presence of a characteristic Rossmann-like fold and the active site sequence DxDx(V ⁄ T). HAD
phosphatases employ an aspartate residue as a nucleophile in a magnesium-dependent phosphoaspartyl transferase reaction. Although there is
genetic evidence demonstrating a causal involvement of some HAD phosphatases in diseases such as cancer, cardiovascular, metabolic and neurological disorders, the physiological roles of many of these enzymes are still
poorly understood. In this review, we discuss the structure and evolution
of human HAD phosphatases, and summarize their known functions in
health and disease.
doi:10.1111/j.1742-4658.2012.08633.x
Introduction
The HAD superfamily is a large and ubiquitous class
of enzymes present in the proteomes of organisms
from all three superkingdoms of life [1]. Over 19 000
unique sequences of superfamily members have been
identified so far, including 183 in Homo sapiens [2]. While
originally named after the haloacid dehalogenases that
Abbreviations
AAA, ATPase associated with diverse cellular activities; BMP, bone morphogenetic protein; BOS, branchio-otic syndrome; BOR, branchiooto-renal syndrome; BRCT, breast cancer protein-related carboxy-terminal; CCFDN, congenital cataracts facial dysmorphism neuropathy
syndrome; CDG1A, congenital disorder of glycosylation type IA; cdN, cytosolic 5‘(3‘)-deoxyribonucleotidase; CLIP, C-terminal lipin domain;
CMD1J, dilated cardiomyopathy type 1J; cN, cytosolic 5‘-nucleotidase; CTD, C-terminal domain; DFNA10, deafness autosomal-dominant
nonsyndromic sensorineural 10 locus; EIEE10, early infantile epileptic encephalopathy-10; eNT, ecto-5‘-nucleotidase; Eya, Eyes absent; Fcp,
TFIIF-associating component of RNA polymerase II CTD phosphatase; FHA, forkhead-associated; GWAS, genome-wide association study;
HAD, haloacid dehalogenase; H. s., Homo sapiens; mdN, mitochondrial 5‘(3‘)-deoxyribonucleotidase; M. m., Mus musculus; NLIP, N-terminal
lipin domain; OMIM, Online Mendelian Inheritance in Man; PDB, protein data bank; Pdxp, pyridoxal 5‘-phosphate phosphatase; PGP,
phosphoglycolate phosphatase; PHGDH, 3-phosphoglycerate dehydrogenase; PMM, phosphomannomutase; PNKP, polynucleotide 5‘-kinase
3‘-phosphatase; PPM, serine ⁄ threonine-directed phosphatase, metal-dependent; PPP, serine ⁄ threonine-directed phosphoprotein
phosphatase; pRb, retinoblastoma protein; PSAT, phosphoserine aminotransferase; PSPH, phosphoserine phosphatase; PTP, protein tyrosine
phosphatase; sEH2, soluble epoxide hydrolase; Six, sine oculis homeobox; Scp, small C-terminal domain phosphatase;
UBLCP1, ubiquitin-like CTD phosphatase.
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Human HAD phosphatases
A. Seifried et al.
are mostly found in prokaryotes, HAD superfamily
members in all organisms are quantitatively dominated
by enzymes that catalyze phosphoryl transfer. The
majority of these phosphotransferases are phosphatases (phosphate monoester hydrolases, 79%) and
ATPases (phosphoanhydride hydrolases, 20%) [2,3].
This review focuses on mammalian HAD-type phosphatases.
Although the overall sequence identity between
HAD phosphatases is typically very low (often
< 15%), family members can be identified by amino
acid sequence alignments based on the presence of four
short HAD signature motifs that contain the conserved
catalytic residues.
HAD phosphatases carry out catalysis differently
from other well-known phosphatases. In contrast to
enzymes from the alkaline phosphatase or tyrosine-specific phosphatase superfamilies that catalyze phosphoryl transfer using serine or cysteine nucleophiles,
respectively, HAD phosphatases use an aspartate residue in the active site for nucleophilic attack [4–7]. This
distinguishing feature of HAD phosphatases also
explains their lack of sensitivity against commonly
employed phosphatase inhibitors and may have contributed to the relatively slow appreciation of their
multiple roles in mammalian cells.
Interest in mammalian HAD phosphoprotein phosphatases has soared with the seminal discovery that
members of the Fcp ⁄ Scp subfamilies act as key regulators of transcription by dephosphorylating the C-terminal domain of RNA polymerase II [8,9], and with
the exciting finding that the Eyes absent (Eya) family
of transcription factors contain intrinsic HAD-type
phosphatase activity that is crucial for organ formation
[10–12]. Due to their phosphotyrosine or phosphoserine
phosphatase activity, Eya or Fcp ⁄ Scp phosphatases
have initially been grouped under phosphotyrosine
phosphatases (PTPs) or metal-dependent serine ⁄
threonine-directed phosphatases [4,13]. However, it is
clear that mammalian HAD phosphatases constitute a
much larger family of enzymes, and that they have
evolved independently from classic phosphatases [14].
Structural and mechanistic features of
HAD phosphatases
modified Rossmann fold, which is characterized by
a three stacked a ⁄ b sandwich, comprised of repeating
b-a units. The central sheet is parallel and generally
consists of at least five strands sequentially arranged in
a ‘54123’ order, thereby orienting four loops which
contain the core residues involved in positioning the
substrate, the cofactor and the catalytic groups
(Fig. 1A, B). The Rossmannoid fold typical of HAD
phosphatases harbors three additional structural signatures that allow the enzyme to adopt distinct conformational states and that contribute to substrate
specificity: the squiggle, flap, and cap domains (discussed below) [2,14,17].
Catalytic mechanism
The characteristic feature of HAD-type phosphatases
is a two-step phosphoaspartyl transferase mechanism.
In the first step, the Asp nucleophile initiates a nucleophilic attack on the phosphoryl group of the substrate,
which results in the formation of a phosphoaspartyl
enzyme intermediate and the displacement of the substrate leaving group. In the subsequent step, a water
molecule exerts a nucleophilic attack on the phosphoaspartyl intermediate, thus releasing free phosphate
and regenerating the catalytic Asp (Fig. 2). HAD
phosphatases contain a second Asp residue positioned
two residues C-terminal of the Asp nucleophile (designated Asp + 2). Asp + 2 functions as a general acid ⁄
base to protonate the leaving group in the first partial
reaction, and to deprotonate the water nucleophile in
the second partial reaction. The chemical advantage of
a catalytic Asp is its versatility: it constitutes both a
good nucleophile and a good leaving group, and it can
operate at low and high pH. All HAD phosphoaspartyl transferases use Mg2+ as an obligatory cofactor.
Mg2+ aids in the correct positioning of the substrate
phosphoryl group relative to the Asp nucleophile, and
electrostatically stabilizes the required close approximation of the anionic nucleophile to the dianionic substrate phosphomonoester (Fig. 1B). Furthermore,
Mg2+ provides charge neutralization of the transition
state. Together, the catalytic residues and the Mg2+
cofactor stabilize the trigonal bipyramidal transition
state of both partial reactions [2].
Rossmannoid fold
Squiggle and flap elements
Crystallographic work on pro- and eukaryotic HAD
phosphatases has revealed that all members of the
HAD phosphatase superfamily share the same structural arrangement of the active core [15,16]. The
residues of the catalytic machinery are positioned in a
The split phosphoaspartyl transferase mechanism is
dependent on an initial reaction that requires solvent
exclusion (to favor the Asp-based nucleophilic attack),
and a subsequent reaction that involves extensive
solvent contact (leading to the hydrolysis of the aspar-
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A. Seifried et al.
A
Human HAD phosphatases
B
Fig. 1. (A) The Rossmann-like fold of the catalytic core of Pdxp ⁄ Chronophin (adapted PDB: 2P69, modified). The structure is reduced to the
residues of the catalytic core. The Rossman fold of HAD phosphatases is formed by repeating b-a units. The central sheet consists of five
b-strands sequentially arranged in a ‘54123’ order (green). The two aspartates of HAD motif I are depicted in red, followed by the squiggle
and flap domains in the first loop (shown in dark gray). (B) Illustration of the steric orientation of the catalytic core residues in the four active
site loops of the human mitochondrial deoxyribonucleotidase (adapted PDB: 1MH9, modified). Nitrogen atoms are blue, oxygen atoms are
red, phosphorus atom is orange, and carbon atoms are gray.
Fig. 2. The general catalytic mechanism of
HAD phosphatases. Catalysis proceeds
through an aspartylphosphate intermediate.
The Asp nucleophile is shown in red,
Asp + 2 is depicted in blue. For details, see
text.
tylphosphate intermediate) [18]. Therefore, an essential
aspect of catalysis is the alternation between closed
and open states of the active site cavity. The basic
structural features in the HAD Rossmannoid core
responsible for this mobility are the unique and con-
served ‘squiggle’ and ‘flap’ signature elements, which
are located immediately downstream of the b1-strand
of the core Rossmannoid fold (see Fig. 1A). The small
squiggle domain of approximately six amino acids
folds into an almost complete single helical turn, while
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A. Seifried et al.
the flap that is located C-terminally of the squiggle
adopts a b-hairpin turn, thus projecting two strands
out from the Rossmannoid core scaffold. The helical
squiggle can switch between tightly or loosely wound
conformations, thereby triggering a movement of the
flap [14]. Since the b-hairpin flap is located immediately adjacent to the active site, this squiggle-induced
flap movement can partly cover the catalytic cavity.
The conformational changes exerted by the squiggle
and flap domains appear to constitute the minimal
machinery required for solvent exclusion and solvent
access at the active site.
Cap modules
Additional mobile inserts termed cap modules can provide more extensive shielding for the catalytic cavity
than the simple flap elements. In addition, cap
domains supply binding determinants for substrate
selectivity, and they can also be involved in phosphatase oligomerization. Despite their structural diversity,
caps can be divided into three broad categories, C0,
C1 and C2, and based on this domain organization,
HAD phosphatases fall within three structural subfamilies (see Table 1).
Whereas C0 elements are very small, C1 and C2 caps
fold into domains of considerable size that move extensively to mediate active site solvent occlusion ⁄ inclusion
during the catalytic cycle (Fig. 3). Type C0 and C1
structures are inserted in between the two b-strands of
the flap itself, whereas C2 caps are incorporated in the
linker immediately after the b3 strand of the core
domain. C0 caps represent the structurally simplest
C0 cap
Polynucleotide 5’-kinase 3’-phosphatase
(PDB: 3ZVL )
modules [19,20], and can consist of loops or b-strands,
as for example in the polynucleotide kinase ⁄ phosphatase (PNKP, PDB: 3ZVL; [21]). The more elaborate
and most common C1 caps are large enough to completely seal the enzyme’s active site in the closed state.
C1 modules in HAD phosphatases are a-helical
domains of varying complexities. For example, a tetrahelical bundle is found in the phosphoserine phosphatase (PSPH, PDB: 1L8L; [16]), and the C1 cap of
Eyes absent 2 (Eya2) (PDB: 3GEB, 3HB0, 3HB1;
[22]) folds into a large bundle of seven helices. The
C2 caps are highly diversified modules, generally composed of a + b domains with a core b-sheet of at
least three strands, to which other simple secondary
structure elements can be added. Examples of C2capped HAD phosphatases include pyridoxal 5¢-phosphatase (Pdxp) ⁄ chronophin (PDB: 2OYC, 2P69; [23])
and phosphomannomutase 1 (PMM1, PDB: 2FUE).
Role of cap modules for substrate selectivity
Due to the location of the catalytic residues at the
C-termini of the b-strands in the Rossmann core (see
Fig. 1), the active site of HAD phosphatases is open.
Therefore, C0 members such as PNKP or the RNA
polymerase II C-terminal domain (CTD) phosphatases
tend to process macromolecular substrates, and the
bound substrate itself functions as a cap by excluding
bulk solvent. In these cases, substrate selectivity may
be provided by a number of invariant residues that line
the entrance to the active site. However, structures of
bacterial HAD phosphatases indicate that uncapped
phosphatases may also utilize small substrates due to
C1 cap
Phosphoserine phosphatase
(PDB:1L8L)
C2 cap
Phosphomannomutase 1
(PDB: 2FUE)
Fig. 3. Structural diversity of HAD phosphatase cap domains. Shown are X-ray crystal structures of PNKP (with an unstructured loop as C0
cap), PSPH (with a C1 cap consisting of a four-helical bundle), and PMM1 (with an a ⁄ b fold as a C2 cap). The catalytic domain is shown in
grey, and cap domains are colored in orange. The catalytic Asp and Asp + 2 are highlighted in red.
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A. Seifried et al.
‘pseudocapping’ by oligomerization via the flap segment [24]. The presence of distinct C1 ⁄ C2-type cap
modules (or the pseudocapping by oligomerization)
sterically restricts access to the catalytic cavity, and
allows phosphatases to act on small molecules, which
can be sequestered within the active site by cap closure. As an exception to this general rule, two capped
phosphatases have been shown to act on macromolecular substrates: Eya, which can dephosphorylate the
C-terminal tyrosyl residue of histone H2AX [22], and
chronophin, which dephosphorylates not only pyridoxal 5¢-phosphate, but also pSer3 of the actin-binding
factor cofilin [23,25]. Therefore, capped proteins can
be accessible to the termini of phosphoproteins.
Structural analysis of C1- and C2-type phosphatases has revealed the existence of ‘substrate specificity
domains’ inserted in the caps [2,26]. These specificity
modules generally consist of residues that interact
with the substrate leaving group, define the electrostatic environment of the active site and activate the
substrate for nucleophilic attack [26,27]. In addition,
substrate binding may also stabilize the closed conformation of the cap domain, thereby providing specificity. As the number of structurally characterized
mammalian HADs increases, it might become possible to identify conserved residues that are responsible
for binding of particular substrate classes. However,
determinants other than specific amino acid residues
strategically placed in the cap modules may be
important for specificity, such as those determining
conformational flexibility and active-site sequestration
[28].
HAD signature motifs
The catalytic core residues are highly conserved
throughout the HAD phosphatase family and cluster
into four signature motifs in the primary amino acid
sequence that correspond to the four active site loop
residues [1,3]. Therefore, the presence of these motifs
provides a means of identifying family members via
amino acid sequence alignments. Based on extensive
work performed mostly on prokaryotes [2,14], HAD
signature motif I contains the essential Asp nucleophile
and has the extended consensus sequence hhhDxDx
(T ⁄ V)(L ⁄ V)h (where h represents a hydrophobic residue, and x indicates any amino acid). The carboxylate
group of the Asp nucleophile and the carbonyl backbone of the second Asp (Asp + 2) in motif I coordinate
the essential Mg2+ in the active site. Motif II with the
consensus sequence hhhhhh(S ⁄ T) contains a conserved
Ser or Thr residue that helps to orient the substrate for
nucleophilic attack by forming a hydrogen bond with
Human HAD phosphatases
its transferring phosphoryl group. Motif III is poorly
conserved in comparison to the other motifs, and centers on a conserved Lys residue, which is spaced 18–30
residues apart from motif IV. The function of the
motif III Lys is to stabilize the negative charge of the
reaction intermediate together with Ser ⁄ Thr of motif II.
Motif IV typically exhibits the consensus sequence
(G ⁄ S)(D ⁄ S)x3-4(D ⁄ E)hhhh, but a DD signature instead
of a Dx3-4D sequence is also observed [29,30]. Together
with the Asp residues of motif I, the conserved motif
IV acidic Asp or Glu residues are involved in the coordination of Mg2+. Motifs I–IV are spatially arranged
around a single binding cavity at the C-terminal end of
the strands of the central sheet that forms the active
site of HAD phosphatases (see Fig. 1A,B).
The gene complement of human HAD
phosphatases
On the basis of the above-discussed criteria that define
the HAD family of phosphatases by the presence of a
Rossmannoid structure of the catalytic core domain
and the active site signature DxDx(V ⁄ T), we have determined the human gene complement of HAD phosphatase catalytic subunits. By database mining, we have
identified 40 different genes and their corresponding
protein products. Fig. 4A shows an alignment of motif I
of 40 human HAD phosphatases and the consensus
motif derived from this alignment, hhhDxDx
(T ⁄ V)(L ⁄ I)h. The family conservation in the 14 amino
acids surrounding motif I is shown in the sequence
logo in Fig. 4B [31]. Figure 4C shows motif II-IV catalytic core residues for selected human HAD phosphatases. In these proteins, we were able to unambiguously
identify the active site residues by inspection of available crystal structures (see legend to Fig. 4C).
Using the Genome Reference Consortium Homo
sapiens high coverage assembly GRCh37 (GRCh37e66,
release February 2012), available via the Ensembl
database, we have also performed a search for the existence of possible HAD phosphatase transcript variants.
As detailed in the Supplementary Table S1, the listed
40 human HAD phosphatase genes can encode for 193
protein-coding transcripts. Thirty of these transcripts
are predicted to be subject to nonsense-mediated
decay, and two transcripts encode for a ‘protein’ product of only two amino acids. Thus, the listed 40
human HAD phosphatases could potentially generate
161 protein-coding phosphatase variants. It remains to
be tested how many of these variants encode functional enzymes. An overview of currently functionally
characterized human HAD phosphatases and their
variants is given in Table 1.
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553
Human HAD phosphatases
Motif
B
I
FE
V
S
SH
AR
Y
H
T
R
DT
W
F
A
G
D
V
S
P
P
D
T
K
I
A
S
H
K
E
LV
L
C
W
C
14
N
A
13
Q
10
3
2
1
S
N
H
C
Motif
VI
Y
M
A
G
F
9
I
F
C
T
8
M
T
7
A
C
DD
V
I
Y
GV
I
A
YA
C
Q
G
S
H
M
6
C
R
L
0
TL
VVF L G
E
VFLW
S
I L
L
I
1
12
2
4
Bits
3
11
4
5
A
A. Seifried et al.
MEME (no SSC) 18.05.12 16:51
Motif
Motif
D
CECR5
CTDNEP CTDP1
ENOPH1 EPHX2
EYA
Human
Homo sapiens
Mammalia
CTDSP CTDSPL2
Vertebrata
HDHD1 LPIN
2
Mouse
Mus musculus
2
3
MDP1
NANP
NT5
NT5D
PGP
PHOSPHO PMM
PSPH
UBLCP1
2
Chordata
NT5C1 NT5C3
3
PNKP
2
Deuterostomia
Zebra fish
Danio rerio
Tunicate
Ciona intestinalis
2
Bilateria
Sea urchin
Strongylocentrotus purpuratus
Ophistokonta
2
Ecdyozoa
2
6
Eumetazoa
4 2
Fruit fly
Drosophila melanogaster
Nematode
Caenorhabditis elegans
Sea anemone
Nematostella vectensis
Placozoa
Trichoplax adherens
2
Sponge
Amphimedon queenslandica
Choanoflagellate
Monosiga brevicollis
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A. Seifried et al.
Considering the existence of 103 genes encoding for
catalytic subunits of Cys-based human PTPs, 15 genes
encoding for catalytic subunits of the PPP family, and
16 genes encoding for human PPM catalytic subunits
[4], HAD phosphatases amount to at least 22.98% of
all human phosphatase catalytic subunits. However,
whereas PPMs (represented by PP2C and pyruvate
dehydrogenase phosphatase) contain catalytic and regulatory domains on one polypeptide chain [7], the catalytic subunits of PPP family members (including PP1
and PP2A as the most abundant Ser ⁄ Thr phosphatases) combinatorially associate with a diverse array of
non-catalytic regulatory and structural proteins,
thereby generating hundreds of functionally distinct
proteins [32]. It is currently not known whether the
catalytic subunits of human HAD phosphatases also
associate with regulatory or targeting subunits. Therefore, the relative abundance of functionally distinct
HAD phosphatases compared to phosphatases containing PTP, PPP, or PPM catalytic subunits can at
present not be precisely assessed.
Evolutionary history of metazoan HAD
phosphatases
The HADs are not only an extremely large, but also a
very old superfamily. As an estimation, five genes
encoding HADs were already present in the last
universal common ancestor [14]. Radiation of these
ur-genes has happened in all three superkingdoms of
life, resulting in at least 23 protein families. Independently, some of these proteins evolved the capability to
delete a phosphate group from a substrate, they
became phosphatases. As most of the radiation happened in bacteria, we asked whether these HAD phosphatases have been a target of evolution also in the
eukaryotic kingdom. Therefore, we traced the evolution of 22 HAD phosphatase families in the kingdom
arguably most relevant for humans, the metazoans
(detailed methods are provided in the Supplementary
Human HAD phosphatases
Text, Doc. S1). We found that most of these families
have already been present in the last common ancestor
of all animals. From here on, different branches have
seen expansion and losses of different families
(Fig. 4D). Most notable are the multiple duplications
in six families at the base of the vertebrates. This
expansion was coupled with the evolution of new functions like in the case of phosphoglycolate phosphatase
(PGP) ⁄ Pdxp and PMM1 ⁄ 2 [33,34]. In addition to these
duplication-driven neofunctionalisations, there seems
to be one case of ‘de novo’ evolution of a phosphatase.
Although present in all analyzed genomes, only the
vertebrate members of the soluble epoxide hydrolase
(sEH2) phosphatases show the hallmark DxDx(V ⁄ T)
motif. Obviously, this sequence-based prediction needs
further experimental characterization. Many evolutionary events also happened in the lineages leading to the
classical model organism Drosophila melanogaster and
Caenorhabditis elegans. Both have seen expansion and
losses affecting six families. Surprisingly, in both
D. melanogaster and C. elegans, the PGP ⁄ Pdxp family
was expanded independently. This coarse-grained analysis reveals that HAD phosphatases are an active target of evolution in metazoans. Thus, the HADs are
not only a useful model to study protein evolution on
the level of superfamilies [14,35], but their enormous
evolutionary flexibility also makes them good candidates to analyze how evolution generates functional
diversity within one protein family.
Multidomain architecture of human HAD
phosphatases
HAD phosphatases have undergone a remarkable
expansion during the evolution of animals (see Fig. 4).
Gene duplication events are typical of higher eukaryotes, and are often accompanied by the acquisition of
new domains which further diversify and specialize
protein functions. Thus, whereas many prokaryotic
HAD phosphatases are small proteins that appear to
Fig. 4. (A) Amino acid sequence alignment of HAD motif I of 40 human HAD phosphatases. The catalytic Asp and Asp + 2 are highlighted
in magenta. (B) Sequence logo depicting the conservation of motif I residues in human HAD phosphatases. MEME suite was used as a
motif-based sequence analysis tool. (C) Amino acid sequence alignments of HAD motifs II–IV of selected human HAD phosphatases. The
highlighted catalytic core residues were identified using the following crystal structures (the PDB code is given in parentheses): ENOPH1
(1YNS), NT5C2 (2J2C), NT5C (2JAR), NT5M (2JAW), PSPH (1L8L), EYA2 (3HB0, 3HB1), PDXP (2P69). The CTDP1 motifs were determined
by alignment with CTDSP1 (1T9Z). (D) Evolution of 22 HAD phosphatase families in metazoa. Blue squares indicate duplication, red squares
loss events. In case of multiple events affecting one family, the number of events is indicated in the squares. Families already present in
Monosiga brevicollis are indicated in black, families emerging later in green. More likely than a family emerging later, it represents a loss in
the more basal genomes like in the case of the NT5C ⁄ D proteins. In the following cases, sequences were grouped together: CTDSP:
CTDSP1, CTDSP2, CTDSPL; EYA: EYA1-4; LPIN: LPIN1-3; MDP1: MDP1, NEDD8-MDP1; NT5: NT5C, NT5D; NT5C1: NT5C1A, NT5C1B;
NT5C3: NT5C3, NT5C3L; NT5D: NT5DC1-4, NT5C2; PGP: PGP, PDXP; PHOSPHO: PHOSPHO1-2; PMM: PMM1-2. For abbreviations, see
Table 1. The detailed methods can be found in the Supplementary Information.
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RNA polymerase C-terminal domain phosphatases
C0
C0
MGDP-1
C0
TFIIF
C0
Scp1
Scp2
Scp3
Fcp1a
Polynucleotide kinase
phosphatase
C0
BRCT
Magnesium-dependent
phosphatase
Intracellular
5`-nucleotidases
C1
cdN
mdN
P5N-I
sEH2
cN-IA
cN-IB
cN-II
Eyes absent
NagD
Cof hydrolases
C2
C2
Chronophin
LHPP
PMM1, 2
C1
Eya1
Eya2
Eya3
Eya4
C0
C1
HAD-phosphatase domain,
C0 cap
C2
HAD-phosphatase domain,
C1 cap
HAD-like hydrolase domain
containing proteins
C1
C1
PNKP
ED2
UBLCP1
CTDNEP1
Soluble epoxide hydrolase
K
C0
HAD-phosphatase domain,
C2 cap
HAD-phosphatase domain,
cap structure unknown
Kinase
domain
HDHD1
HDHD4
Lipins
NLIP
S/S/S
Lipin1
Lipin2
Lipin3
Forkheadassociated domain
K
PSPH
PHOSPHO1
PHOSPHO2
Epoxide
hydrolase
Transactivation
domain
Mitochondrial leader sequence
(mdN only)
BRCT
S/S/S region of regulatory serines
Breast cancer
TranscriptionTFIIF Factor interacting
Protein-related
Carboxy-terminal
Helix
Transmembrane
Helix motif
P/S/T
Rich region
NLIP
Amphipathic
domain
Ubiquitin-like domain
Fig. 5. Schematic view of the domain structures of characterized human HAD phosphatases. For further details, see text. *These proteins
belong to different HAD families (PSPH, PHOSPHO1 ⁄ 2: pyrimidine 5¢-nucleotidase ⁄ phosphoserine phosphatases) and HDHD1 ⁄ 4 (HAD-like
hydrolase domain-containing proteins), but are grouped here due to their common C1 cap and the absence of other recognizable functional
domains.
consist of a single hydrolase domain, some human
HAD phosphatases contain additional domains that
give some indications as to their subcellular localization and functions (Fig. 5).
While there are no known examples of HAD phosphatases with extracellular domains (as can be found
in receptor PTPs), the C-terminal domain nuclear
envelope phosphatase ⁄ dullard homolog has a transmembrane helix motif required for nuclear membrane
targeting [36], and the mitochondrial deoxyribonucleotidase mdNT contains a mitochondrial leader sequence
[26]. The specialized functions of PNKP and sEH2 for
DNA repair or lipid metabolism, respectively, have
been accomplished by the fusion of HAD phosphatase
domains with DNA kinase or epoxide hydrolase
domains [21,37]. PNKP additionally contains a DNA
556
binding motif and a forkhead-associated domain that
mediates binding to other DNA repair proteins. The
Eya phosphatase domain is embedded in a region that
mediates protein-protein interactions with DNA binding proteins, and the catalytic domain is additionally
fused to a transactivation domain flanked by P ⁄ S ⁄ Trich regions [38]. To fulfill their functions in lipid
metabolism on intracellular membranes, lipin phosphatases contain an amphipathic a-helix responsible for
membrane association. In addition, lipins contain a
nuclear localization signal and coactivator motifs to
regulate the transcription of genes involved in fatty
acid metabolism [39]. The RNA polymerase II
C-terminal domain (CTD) phosphatase Fcp1 contains
a transcription factor TFIIF-interacting helix and a
breast cancer protein-related carboxy-terminal (BRCT)
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domain that binds to the phosphorylated CTD [20,40],
whereas the Fcp1-related small CTD phosphatases
(Scps) lack the BRCT and TFIIF-binding domains
[19], and the ubiquitin-like CTD phosphatase (UBLCP1) is additionally equipped with an ubiquitin-like
domain [41].
Thus, while some HAD phosphatases have an
elaborate extracatalytic multidomain structure, others
display no additional recognizable domains. These
phosphatases may associate with regulatory or targeting subunits (although very few interacting proteins of HAD phosphatases have been described so
far), or they may operate as single hydrolase domain
entities whose specificity is determined by their cap
structure.
We have also performed a preliminary bioinformatic
exploration of the link between the combination of
multiple biochemical activities in some HAD phosphatases and the evolution of these enzymes. Although
many of the domains found in eukaryotic HADs have
already been present in prokaryotes, their fusion is
eukaryote specific. The only exception is the combination of the HAD domain with a kinase domain of the
AAA family in PNKP, which is also present in bacteria. As here the order of the domains is inversed, this
fusion seems to have happened at least twice independently. The phylogenetic distribution of the domain
architectures of lipins, PNKP and UBLCP1 suggest an
origin in the last common ancestor of the eukaryotes.
Also, the core architecture of CTDP1 (HAD +
BRCT) was identified throughout all eukaryotes. The
additional accretion of the TFIIF domain happened at
the base of the vertebrates with the exception of one
protein in the beetle Tribolium castaneum. The most
recent event was the fusion of HAD with the epoxide
hydrolase domain at the base of the tetrapods. Thus,
the enzymatic combinations found in human multidomain HAD phosphatases serve to further diversify
and specialize HAD phosphatase functions.
Roles in human health and disease
A number of HAD phosphatases play important roles
in a range of human diseases, including cancer, cardiovascular, metabolic and neurological disorders (see
Table 1). This overview chapter highlights those HAD
phosphatases whose causal link to human disease is
supported by genetic or epidemiological data.
Whenever applicable, Table 1 also contains references to the Online Mendelian Inheritance in Man
(OMIM) database, a genetic database that curates the
medical literature for genetic disorders [42]. In cases in
which individual genes have been associated with a
Human HAD phosphatases
physiological phenotype, OMIM provides clinical
descriptions together with some genetic information.
However, the listed genetic variants may be linked to a
biological phenotype more by statistical association
than necessarily by functional or medical analysis. It
has to be taken into account that OMIM does not list
all known variants for each gene, and that OMIM
does not attempt to report all genes and variants identified through genome-wide association studies
(GWAS). For an in-depth analysis of all variants identified for a particular gene, the reader should therefore
consult additional databases, such as GWAS central
(https://www.gwascentral.org), HGMD (http://www.
hgmd.org), LOVD (http://www.lovd.nl/2.0/), or MutaDATABASE (http://www.mutadatabase.org/) [43].
RNA polymerase C-terminal domain phosphatase
(CTDP1 ⁄ Fcp1)
The unstructured C-terminal domain (CTD) of eukaryotic RNA polymerase II regulates transcription by
recruiting different factors to nascent mRNA. The
human CTD is composed of 52 tandem heptapeptide
repeats with the sequence Y1S2P3T4S5P6S7, which are
dynamically phosphorylated and dephosphorylated
throughout transcription cycles. The extent and pattern of CTD phosphorylation represents a critical regulatory checkpoint for transcription and is determined
by dedicated CTD kinases and phosphatases. Transcription initiation requires CTD dephosphorylation
by phosphatases, which are therefore essential for the
regulation of gene expression [20].
Fcp1 is the main serine phosphatase for the CTD
and can processively dephosphorylate both pSer2 and
pSer5. Varon et al. [44] have shown that the congenital cataracts facial dysmorphism neuropathy syndrome (CCFDN; OMIM #604168) is caused by Fcp1
loss-of-function. A single-nucleotide substitution in an
antisense Alu element in intron 6 of CTDP1 (encoding for Fcp1) results in a rare mechanism of aberrant
splicing and an Alu insertion in the processed
mRNA. The insertion in the CTDP1 mRNA results
in a premature termination signal 17 codons downstream of exon 6, with the mutant transcript expected
to undergo nonsense-mediated decay or to produce a
nonfunctional protein lacking the nuclear localization
signal.
CCFDN is an autosomal recessive disorder prevalent among Gypsy families. This demyelinating neuropathy is characterized by progressive peripheral
nerve abnormalities that lead to severe disability. It is
currently not understood how nonfunctional Fcp1a
results in the specific symptoms of CCFDN.
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Small C-terminal domain phosphatases (Scp1-3)
Scps are structurally related to Fcp1, and control the
RNA polymerase II transcription machinery by preferentially dephosphorylating the CTD on pSer5 [8]. The
expression of Scp1-3 is confined to non-neuronal tissues and neuroepithelial precursor cells, where they
operate in a silencing complex to epigenetically block
the inappropriate expression of specific neuronal genes
[45]. Since antagonism of the Scp pathway might promote neuronal stem cell differentiation in vivo, small
molecule Scp phosphatase inhibitors could be powerful
tools to direct neurogenesis and promote the regeneration of neurons, for example upon neuronal injury.
Zhang and colleagues have targeted the unique hydrophobic binding pocket adjacent to the Scp active site
[19,46], and have identified rabeprazol as a first lead
compound that selectively inhibits Scp1, but not the
related Fcp1 or Dullard proteins [47]. This study provides a promising starting point for the design and
optimization of potent and specific Scp inhibitors
that may facilitate neuronal differentiation to repair
nervous system damage.
Besides the regulation of transcription by CTD
dephosphorylation, Scps can also recognize other substrates and fulfil additional biological functions.
Scps1-3 can dephosphorylate and stabilize Snail, a key
transcriptional repressor of E-cadherin. Stabilization of
Snail by Scp enhances E-cadherin promoter suppression and promotes cell migration in vitro [48]. Scps1-3
also dephosphorylate and modulate the activities of
Smad1 and Smad2 ⁄ 3 proteins, which function as critical transducers of bone morphogenetic protein- and
transforming growth factor b-initiated cellular
responses [49,50]. Transiently overexpressed Scp3 can
dephosphorylate the tumor suppressor retinoblastoma
protein 1 (pRb1) in cells, and may thereby activate
Rb1 to inhibit cell cycle progression [51]. This finding
may explain a role of Scp3 in cancer: CTDSPL
(encoding for Scp3) resides in a chromosomal region
(3p21.3) that is deleted in > 90% of major human
carcinomas, including small cell lung cancer, renal cell
carcinoma and breast carcinoma. Scp3 was found to
be hemi- or homozygously deleted or functionally inactivated by mutations in some of these malignancies,
and was additionally shown to function as a tumor
suppressor in immunocompromised mice [51].
Polynucleotide 5¢-kinase ⁄ 3¢-phosphatase (PNKP)
DNA damage occurs constantly by various internal
agents (such as reactive oxygen species), during normal
processes such as DNA replication, and by external
558
agents (such as ultraviolet light). DNA damage is continuously repaired by several DNA repair pathways
[52,53], and defects in these processes are considered to
play a causative role in aging [54] and neurological disorders [55], and to be an important factor in the etiology and treatment of cancer [56]. Ionizing radiation
and other internal and external DNA damaging agents
often generate DNA strand breaks with incompatible
termini, which first require processing before strand
resynthesis and ligation by DNA polymerases and
ligases can take place. Termini with 3¢-phosphate and
5¢-hydroxyl groups occur very frequently, and PNKP
is the major enzyme that restores the chemistry of
strand breaks by generating the obligatory 3¢-hydroxyl
and 5¢-phosphate termini for repair [53].
PNKP is a multidomain enzyme that consists of an
N-terminal forkhead-associated (FHA) domain and a
C-terminal catalytic domain, composed of fused HAD
phosphatase and kinase subdomains [21]. Interestingly,
the phosphatase activity of PNKP appears to be much
higher than its kinase activity, which may reflect the
more frequent occurrence of 3¢-phosphorylated termini
upon DNA damage [57]. PNKP is a key enzyme in
several DNA repair pathways (i.e., single-strand break
repair, base-excision repair and double-strand break
repair), because it interacts with other DNA repair
proteins, notably with phosphorylated XRCC1 and
XRCC4, via its FHA domain [53].
Genetic defects in PNKP have revealed its essential
role in the developing central nervous system. Neurons
are particularly sensitive to mutations in DNA repair
genes, and loss-of-function mutations in PNKP (leading to substitutions in the kinase and phosphatase
domains) cause early infantile epileptic encephalopathy-10 (EIEE10; OMIM #613402) [58]. EIEE10 is a
severe, autosomal recessive disease that is characterized
by intractable seizures, microcephaly and developmental delay.
In cancer cells, on the other hand, PNKP-mediated
DNA repair can enhance the resistance to genotoxic
therapeutic agents. The DNA repair capacity of tumor
cells is regarded as an important factor in the clinical
response to ionizing radiation and various chemotherapeutic agents, because it protects cells from genotoxic
insults. Blocking DNA repair sensitizes tumor cells to
apoptosis, and the recent identification of small molecule DNA repair protein inhibitors has increased clinical interest in this pharmacological concept. PNKP
emerges as a particularly attractive therapeutic target
due to its importance in multiple DNA repair pathways. A non-competitive, allosteric small molecule
inhibitor of PNKP phosphatase activity has been identified [59]. This polysubstituted imidopiperidine
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compound specifically blocks human PNKP, but not
other related DNA phosphatases, PP-1cc, or calcineurin. Given this lead structure, new inhibitory compounds will need to be identified and optimized for
clinical use.
Soluble epoxide hydrolase 2 (sEH2)
Endogenous fatty acid epoxides such as the arachidonic acid-derived epoxyeicosatrienoic acids are signaling molecules that possess a wide variety of biological
effects, many of which are related to cardiovascular
physiology and inflammation. The human sEH2 is a
homodimeric, bifunctional enzyme with a C-terminal
epoxide hydrolase domain, which is responsible for the
transformation of epoxyeicosatrienoic acids to the corresponding vicinal diols. sEH2 also has an N-terminal
HAD-type phosphatase domain, whose function has
long remained elusive [37,60]. Recent studies demonstrate that the sEH2 N-terminal domain can effectively
dephosphorylate dihydroxy lipid phosphates and polyisoprenyl pyro- and monophosphates, which are metabolic precursors of cholesterol biosynthesis [61].
sEH2 has been identified as a heart failure susceptibility gene in a model of spontaneously hypertensive
heart failure rats [62]. Monti et al. found increased
sEH2 expression and elevated epoxide hydrolase activity, leading to a more rapid hydrolysis of cardioprotective epoxyeicosatrienoic acids. Furthermore, EPHX2
gene ablation in mice protected from pressure overload-induced heart failure and cardiac arrhythmias.
While a potential contribution of the sEH2 phosphatase activity was not investigated in this study, new
findings indicate that sEH2 phosphatase activity may
play an important pathophysiological role.
First, while the pharmacological inhibition of sEH2
epoxide hydrolase activity in vivo attenuates hypertension, this effect was markedly less impressive than the
blood pressure reduction observed upon complete
EPHX2 gene deletion in mice, pointing to a role of the
phosphatase domain for blood pressure elevation [37].
Second, sEH2 phosphatase activity appears to contribute significantly to the role of sEH2 in lipid metabolism and lipid-related disorders. sEH2 phosphatase
activity leads to an elevation of cholesterol levels,
whereas the sEH2 epoxide hydrolase activity lowers
cholesterol levels in cells and the administration of an
sEH2 epoxide hydrolase inhibitor elevated cholesterol
levels in vivo [61].
Third, several epidemiological studies link human
EPHX2 polymorphisms with dyslipidemia and related
disorders, such as atherosclerosis and coronary heart
disease. The most frequently found EPHX2 SNP leads
Human HAD phosphatases
to a R287Q substitution, which is primarily associated
with cardiovascular disease. Importantly, sEH2-R287Q
displays significantly impaired epoxide hydrolase activity, but elevated phosphatase activity. sEH2-R287Q
can act as a modifier of familial hypercholesterolemia
caused by a heterozygous mutation in the low density
lipoprotein receptor (LDLR) gene, and is associated
with elevated plasma triglyceride levels (OMIM
#143890). Other epidemiological studies have found an
association of the sEH2-R287Q polymorphism with
elevated risk for coronary artery calcification and subclinical arteriosclerosis [63], and EPHX2 haplotypes
have also been associated with altered risk of ischemic
stroke (OMIM #601367).
Based on the potent anti-inflammatory, vasodilator,
and cardioprotective properties of epoxyeicosatrienoic
acids, sEH2 epoxide hydrolase inhibitors are currently
being developed as potential therapeutic strategies for
the treatment of inflammatory disorders and cardiovascular diseases [64,65]. The findings described above
add a cautionary note to the development of sEH2
hydrolase-targeted inhibitors for the treatment of cardiovascular diseases associated with dyslipidemia. Further studies are required to elucidate the mechanism of
the sEH2 phosphatase-dependent increase of cholesterol levels. Ultimately, inhibitors of sEH2 phosphatase activity may offer novel therapeutic approaches
for the management of dyslipidemia-related disorders.
Intracellular 5¢-nucleotidases
The human 5¢-nucleotidases are a large family of genetically unrelated enzymes that catalyze the dephosphorylation of (deoxy)ribonucleoside monophosphates
[(d)NMPs] to the corresponding nucleosides, and function to maintain balanced cellular NTP and dNTP
pools [66–68]. Characterized family members are
ecto-5¢-nucleotidase (eNT), cytosolic 5¢-nucleotidase
(cN)-IA, cN-IB, cN-II, cN-III, cytosolic 5¢(3¢)-deoxyribonucleotidase (cdN) and mitochondrial 5¢(3¢)-deoxyribonucleotidase (mdN). Additional, related sequences
can be identified in databases (see Fig. 4). While eNT
does not harbor typical HAD domains, HAD motif I is
found in all intracellular 5¢-nucleotidases. HAD motifs
II-IV were identified only in mdN, cdN and cN-III. The
crystal structure of human mdN has revealed the presence of a specificity motif (motif S) that forms hydrogen
bonds with the substrate base [26], and motif S is present in all intracellular 5¢-nucleotidases [66].
5¢-Nucleotidases differ in their affinities for
(d)NMPs, their subcellular localization and tissue distribution. The non-HAD-type eNT (also known as
CD73) is an ubiquitous, AMP-hydrolyzing enzyme
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bound to the external leaflet of the plasma membrane.
eNT produces extracellular adenosine, a ligand for G
protein-coupled, purinergic receptors with important
functions in cellular signal transduction. The HAD
phosphatases cN-IA ⁄ -IB are cytosolic nucleotidases,
characterized by their affinity towards AMP, while
cN-II is an IMP and ⁄ or GMP preferring enzyme.
cN-III is a pyrimidine 5¢-nucleotidase, and mdN and
cdN are 5¢(3¢)-pyrimidine nucleotidases. In addition to
their nucleotidase activities, cN-II, cN-III and cdN
have also been demonstrated to act as phosphotransferases, i.e., the phosphate from the phosphoaspartyl
intermediate is transferred to another nucleoside instead
of being hydrolyzed by water. In contrast, cN-IA does
not display phosphotransferase activity [66].
Intracellular HAD-type 5¢-nucleotidases play a general role in the salvage pathway to recover nucleosides
that are formed during RNA and DNA degradation
for the synthesis of nucleotides and in nucleic acid
repair [68,69]. In addition, cN-III is involved in breakdown of pyrimidine nucleotides during erythrocyte
maturation [70]. Aside from these physiological functions, 5¢-nucleotidases appear to play a role in the
development of drug resistance against nucleoside analogues [71]. Nucleoside analogues are important antimetabolites used in the treatment of cancer and viral
infections [72,73]. These drugs inhibit DNA synthesis
either directly, or through inhibition of DNA precursor synthesis by acting on the de novo or salvage pathways. Nucleoside analogues mimic natural nucleosides,
and need to be converted to their active triphosphate
forms in cells. 5¢-(Deoxy)nucleotidases can dephosphorylate and thus inactivate the monophosphate
forms of nucleoside analogues, and clinical data indicate that 5¢-(deoxy)nucleotidases contribute to the
development of nucleoside analogue resistance. Of the
5¢-nucleotidases, cN-II has so far received most attention for its possible role in resistance to antimetabolites. cdN is also a good candidate for mediating
nucleoside analogue resistance, due to its preference
for the deoxyribonucleoside monophosphates that
nucleoside analogues are designed to mimic [66].
Cytosolic 5¢-nucleotidases-IA, -IB
Human cytosolic nucleotidases cN-IA and cN-IB are
two closely related, AMP preferring 5¢-nucleotidases.
cN-IB is poorly characterized, but appears to be functionally similar to cN-IA. cN-IA expression predominates in the heart, where it is found associated with
the contractile elements of cardiomyocytes. The main
function of cN-IA appears to be the intracellular formation of adenosine from AMP under conditions of
560
ATP breakdown, such as ischemia and hypoxia. cN-I
requires a nucleoside diphosphate such as ADP for
maximum activity. Thus, ATP consuming conditions
increase cN-I activator (ADP) and substrate (AMP)
concentrations, which together with decreased adenosine kinase activity ensure sufficient adenosine generation. The produced adenosine is then excreted and
stimulates purinergic adenosine cell surface receptors,
thereby increasing coronary blood flow, antagonizing
the effects of catecholamines, and prolonging atrioventricular conduction time. Together, these actions of
adenosine increase energy supply and reduce energy
demand of the heart. A selective inhibitor of cN-IA
has been developed, which effectively blocks adenosine
formation in rat cardiomyocytes [reviewed in 66].
Cytosolic 5¢-nucleotidase III (cN-III)
cN-III catalyzes the dephosphorylation of the 5¢-pyrimidine monophosphates CMP and UMP to the corresponding nucleosides during RNA degradation in
maturing erythrocytes. Although cN-III is found in various other cells and tissues, its function has mainly been
studied in red blood cells. Deficiency of NT5C3 due to
mutations in the gene causes autosomal recessive, nonspherocytic hemolytic anemia, characterized by a massive accumulation of pyrimidine nucleotides within the
erythrocyte that interfere with glycolysis (OMIM
#266120). After glucose 6-phosphate dehydrogenase
and pyruvate kinase deficiencies, cN-III deficiency is the
third most common cause of a red blood cell enzymopathy causing hemolysis [74].
Phosphoserine phosphatase (PSPH)
l-Serine is a non-essential amino acid that is available
from dietary protein, protein and phospholipid degradation, and biosynthetically. The major endogenous
source is the glycolytic intermediate 3-phosphoglycerate. This biosynthetic pathway involves three enzymes,
3-phosphoglycerate dehydrogenase (PHGDH), phosphoserine aminotransferase 1 (PSAT1) and PSPH that
catalyzes the final and irreversible step. l-Serine is
essential for the synthesis of proteins and other biomolecules needed for cell proliferation, including nucleotides, phosphatidyl-serine and sphingosine. l-Serine is
also a precursor for the neuromodulators d-serine and
glycine, both of which function as endogenous ligands
at the ‘glycine site’ of the N-methyl-d-aspartat receptor
[75], an ionotropic glutamate receptor with important
roles for memory and learning [76].
Patients with congenital defects in the l-serine synthesizing enzymes present with severe neurological
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abnormalities, demonstrating that the de novo synthesis
of l-serine plays an essential role in the development
and functioning of the central nervous system. Among
these, PSPH deficiency syndrome (OMIM #614023) is
an autosomal recessive disorder caused by PSPH polymorphism. The syndrome is characterized by strongly
reduced enzymatic PSPH activity, due to a PSPHD32N substitution (resulting in 50% residual phosphatase activity), and a PSPH-M52T substitution (with no
detectable residual enzymatic activity). Affected individuals present with intrauterine and postnatal growth
retardation, congenital microcephaly, feeding difficulties and moderate psychomotor retardation. Some of
these symptoms can be alleviated by substitution treatment with oral serine [77,78].
The l-serine biosynthetic pathway is important for
cell proliferation, and has therefore been extensively
studied as a potential target in cancer treatment. Via a
negative-selection RNAi screening using a human breast
cancer xenograft model at an orthotopic site in the
mouse, Sabatini and colleagues have recently shown
that the serine synthesis pathway is essential for tumorigenesis in estrogen receptor-negative breast cancer [79].
PHGDH was amplified and PHGDH protein levels
were elevated in the majority of the disease cases, and
RNAi-mediated inhibition of PHGDH, PSAT1 and
PSPH blocked tumor formation. Interestingly, serine
production was not the only important role of PHGDH
in these tumor cells, and the authors showed that the
pathway contributes substantially to the anaplerosis of
glutamate into the tricarboxylic acid cycle by producing
a-ketoglutarate. These findings suggest that inhibitors
of serine biosynthesis may be of value in the treatment
of estrogen receptor-negative breast tumors [79].
A chemical genetic screen has identified P-Ser as an
inhibitor of neural progenitor proliferation that stimulates neurogenic fate commitment, terminal differentiation, and nascent neuronal survival via the activation
of the metabotropic glutamate receptor 4. These results
suggest that elevating P-Ser levels by inhibiting PSPH
may be of therapeutic value, e.g., for the therapy of
stroke or spinal cord injury [80]. A competitive PSPH
inhibitor, AP3, has been described. This compound is
a structural analog of P-Ser and also functions as a
metabotropic glutamate receptor antagonist [81].
Eyes absent (Eya)
Eya proteins belong to a novel family of proteins identified in many animals. Humans have four paralogs,
designated Eya 1-4. Eya proteins have been named
after their critical function in a conserved network of
transcription factors collectively termed the ‘retinal
Human HAD phosphatases
determination gene network’ for their role in Drosophila eye specification. Mammalian Eya proteins are
involved in the formation of many tissues and organs,
and mutations in human Eya proteins cause a variety
of congenital disorders [38,82].
Eya proteins are defined by a highly conserved 270
amino acid C-terminal motif referred to as the Eya
domain. This domain is required for interaction with a
homeodomain protein called Sine oculis in Drosophila
and Six in vertebrates, and with the transcriptional regulator Dachshund, named Dachshund homolog 1 in mice.
Eya and Six operate as a transcription factor complex,
in which Six mediates DNA binding, and Eya uses its
N-terminal domain for the activation of transcription.
The realization that the Eya domain contains
embedded HAD phosphatase signature sequences, and
that Eya proteins are indeed functional phosphatases,
was a breakthrough in developmental biology because
it provided the first example of a transcription factor
with an inherent phosphatase activity [10–12]. This
work also raised awareness of the large and heterogeneous group of HAD phosphatases that had previously been characterized in prokaryotes, but had gone
mostly unnoticed in higher eukaryotes.
While initial work in transfected cells had indicated
that Eya’s phosphatase activity could be pivotal for
the Eya-mediated transcriptional activation of some
Six-dependent reporter genes [11], recent studies in
Drosophila showed that reducing Eya phosphatase
activity does not globally impair transcriptional output
[83]. Eya proteins function as protein tyrosine phosphatases, although the identification of physiological
Eya targets remains an important issue. It has been
demonstrated that Eya3 can dephosphorylate PTyr-142
of histone H2AX, a decisive phosphorylation mark
that discriminates between apoptotic or DNA repair
responses to genotoxic stress [84,85]. The H2AX
dephosphorylation by Eya3 promotes the recruitment
of DNA repair complexes and thus renders cells resistant to apoptosis. Therefore, the phosphatase activity
of Eya may block an improper apoptotic response to
physiological levels of genotoxic stress by dephosphorylating H2AX on tyrosine, and this function may be
critical in mammalian organogenesis. Eya proteins have
also been reported to dephosphorylate threonine residues, but this activity is apparently encoded in the Nterminal portion of the protein. This non-HAD-type
threonine phosphatase activity of Eya4 has recently
been linked to the regulation of antiviral innate immune
responses by modulating the phosphorylation state of
signal transducers for intracellular pathogens [86].
Mutations in the EYA1, SIX1, and SIX5 genes
cause branchio-oto-renal syndrome (BOR1, OMIM
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#113650), and branchio-otic syndrome (BOS1, OMIM
#602588). Mutations in EYA1 are detected in approximately 40% of affected individuals, whereas SIX mutations are much less common. BOR1 is an autosomal
dominant disorder that is characterized by fistulas or
cysts in the neck, hearing loss (found in > 90% of
BOR1 patients), ear malformations and abnormalities
of kidney structure and function, ranging from mild
renal hypoplasia to a complete lack of kidney formation. BOR1 is estimated to affect about 1 in 40 000
people. BOS1 can be caused by allelic variants of
EYA1, and is characterized by branchial and otic
anomalies as seen in individuals with BOR1, in the
absence of renal anomalies. Molecular genetic testing
is clinically available.
A deletion mutation in human EYA4 has been identified as a cause of dilated cardiomyopathy type 1J
(CMD1J) and heart failure, preceded by sensorineural
hearing loss (OMIM #605362). The transmission of
this genetic disorder is autosomal dominant. Biochemical analysis indicated that the shortened peptide of
Eya4 produced by the deletion mutation failed to bind
wildtype Eya4 and Six proteins, suggesting that its
functions as a transcriptional co-activator may be
impaired [87]. Mutations in human EYA4 have also
been identified at the deafness, autosomal dominant
nonsyndromic sensorineural 10 locus (DFNA10;
OMIM #601316). Affected individuals exhibit a postlingual, progressive form of deafness that can finally
lead to severe-to-profound hearing impairment. This
disorder is caused by a truncation that deletes the Eya
domain, but not the variable domain of Eya4 [88].
Since dilated cardiomyopathy has not been observed,
the partial truncation of the Eya4 variable domain
observed in CMD1J correlates with the occurrence of
dilated cardiomyopathy. Because sensorineural hearing
loss is generally caused by abnormalities in the hair
cells of the organ of Corti in the cochlea, the phenotype of individuals affected by CMD1J and DFNA10
indicates that Eya4 is also important postdevelopmentally for the continued function of the mature organ of
Corti.
In Drosophila, Eya and Sine oculis overexpression
triggers tissue overgrowth [82], and elevated levels of
Eya and Six family members have been observed in
some malignant tumors in humans, including breast
and ovarian cancers and malignant peripheral nerve
sheath tumors [89–92]. In ovarian cancer, Eya2 is upregulated on the RNA and protein levels, in part due
to genomic amplification, and this overexpression is
significantly associated with short overall survival [90].
Furthermore, the ectopic expression of Eya2 in xenograft tumors significantly promotes tumor growth
562
in vivo [90]. Conversely, RNA interference-mediated
suppression of EYA4 expression in malignant peripheral nerve sheath tumor cells suppresses tumor growth
in nude mice [89].
Transcriptional targets of mammalian Eya and Six
proteins include not only the cell cycle regulatory genes
cyclin D1 and cyclin A1, but also the proto-oncogene
c-Myc, and ezrin, a regulator of the cytoskeleton and
contributor to cell migration and metastasis [83].
Indeed, both Six1 and Eya have independently been
shown to mediate cancer metastasis [91,93–95], and it
is the tyrosine phosphatase activity of the Eyas that is
essential to promote breast cancer cell migration, invasion, and transformation in vitro [91]. Using RNA
interference-mediated depletion of Eya2 in MCF7
mammary carcinoma cells, it has recently been demonstrated that Six1 and Eya2 functionally interact during
tumor progression, and that Eya2 is a necessary
co-factor for many of the metastasis promoting functions of Six1 [96]. These findings suggest that targeting
the Six1-Eya interaction may represent a novel strategy
to inhibit breast cancer progression.
Phosphomannomutase-2 (PMM2)
PMM2 converts mannose 6-phosphate to mannose 1phosphate, which is then transformed to GDP-mannose [97]. This mannose donor is needed for the initial
step of protein N-glycosylation. Oligosaccharide moieties on glycoproteins can determine their folding,
transport, biological activity and stability. Therefore,
protein glycosylation errors can affect a broad spectrum of cellular functions, including metabolism, cell
recognition, adhesion and migration, host defense and
antigenicity. Defying its namesake phosphomannomutase, the PMM2 paralog PMM1 has recently been
shown to act as an IMP-stimulated glucose-1,6-bisphosphatase that may be involved in brain metabolism
under ischemic conditions [98].
While PMM1 has not been linked with any hereditary disease, mutations in the PMM2 gene cause the
congenital disorder of glycosylation, type Ia (CDG1A,
Jaeken syndrome, PMM2-CDG (CDG-Ia); OMIM
#212065; [99]). CDGs are autosomal recessive disorders, and CDG1A is the most widespread form with
an estimated prevalence as high as 1 : 20 000. In individuals with enzymatically proven CDG1A, the mutation detection rate in PMM2 is as high as 100% and
includes missense mutations and deletions. Molecular
genetic approaches have been established to detect
PMM2 sequence variants and exonic or whole-gene
deletions of the PMM2 locus by sequence analysis or
deletion ⁄ duplication analysis, and these methods are
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available for clinical testing. CDG1A usually presents
as a severe neurological disorder in the neonatal period. The clinical phenotype of CDG1A is broad, with
basic signs including developmental delay, cerebellar
atrophy, peripheral neuropathy, hypotonia and psychomotor retardation. The lethality in the first year of
life is 20% due to severe infections, liver insufficiency,
or cardiomyopathy. There is currently no pharmacological option to correct the glycosylation defect in
CDG1A patients, and a better understanding of the
pathophysiology is needed to enable the development
of therapeutic strategies [100].
Lipins
The triglyceride core of lipid droplets is mainly synthesized through the sequential acylation of glycerol3-phosphate. The penultimate step in triacylglycerol
synthesis consists in the Mg2+-dependent dephosphorylation of phosphatidic acid to form diacylglycerol.
This key step in lipid biosynthesis is catalyzed by a family of HAD-type phosphatases called lipin1-3 [39].
Emerging evidence suggests that lipins also play crucial
roles in the nucleus as transcriptional coactivators that
regulate the expression of genes involved in lipid metabolism. The lipins exhibit distinct patterns of tissuespecific expression and appear to play non-redundant
roles, with lipin1 being principally expressed in adipose
tissue, skeletal muscle, and heart; lipin2 found predominantly in the liver, and lipin3 in the intestine. All three
mammalian lipins possess phosphatidic acid phosphatase activity, but lipin1 is by far the most active
enzyme. Common features of all lipins are their highly
conserved N- and C-terminal lipin domains (termed
NLIP and CLIP). The CLIP domain contains the transcriptional coactivator motif and the HAD domains
required for phosphatidic acid dephosphorylation.
Lipin1-deficiency was identified as the cause of the
disturbed metabolic phenotype of the fatty liver dystrophy (fld) mouse [101]. Fld mice are lipodystrophic,
exhibit multiple defects in adipose tissue development,
and are characterized by insulin resistance, peripheral
neuropathy and neonatal fatty liver. While inactivating
LPIN1 mutations in humans are not associated with
lipodystrophy for reasons that are currently unclear,
human LPIN1 polymorphisms have been linked to
rhabdomyolysis (also known as autosomal recessive
recurrent acute myoglobinuria, OMIM #268200). Disease onset is typically in childhood, and can be fatal
due to generalized muscle weakness and kidney failure.
Using homozygosity mapping, Zeharia et al. [102] have
identified six mutations in the LPIN1 gene in patients
who presented with recurrent, massive rhabdomyolysis.
Human HAD phosphatases
These mutations create stop codons at residues 215,
388, and 800 or produce frame shifts as a result of
exon skipping, and are thus predicted to result in truncated proteins lacking catalytic activity. Consistent
with these molecular findings, analysis of the muscle
tissue phospholipid contents demonstrated an accumulation of the lipin-1 substrates phosphatidic acid and
lysophospholipids. Interestingly, the authors also identified one carrier for a pathogenic mutation in the
LPIN1 gene (Glu769Gly) among six individuals who
developed statin-induced myopathy [102]. Several other
studies have linked multiple additional polymorphisms
in the LPIN1 gene (leading to lipins with impaired catalytic activities) to metabolic disease traits, such as
insulin resistance and diabetes, blood pressure regulation, response to thiazolidinedione drugs, and susceptibility to statin-induced myopathy [101,103–106].
Mutations in the human LPIN2 gene, leading to lipin 2 deficiency, can cause a very rare autoinflammatory bone disease known as Majeed syndrome (OMIM
#609628), which is characterized by recurrent multifocal bone and skin inflammation and dyserythropoietic
anemia. The pathomechanism is currently unclear, but
may be related to an accumulation phosphatidic acid
that may trigger inflammatory signaling cascades.
Conclusions and future perspectives
HAD phosphatases constitute an ancient, large and
very diverse group of enzymes that have evolved to
specifically dephosphorylate carbohydrates, lipids,
metabolites, DNA and serine-, threonine- or tyrosinephosphorylated proteins in humans. While they have
long been regarded as metabolic phosphatases with
relaxed substrate specificities that fulfil merely housekeeping functions, recent findings prove otherwise. It is
now clear that loss of some HAD phosphatases causes
hereditary disorders, and evidence is accumulating that
several human HAD phosphatases are involved in
important diseases, such as cancer, cardiovascular,
metabolic and neurological disorders.
It can be expected that the characterization of the
physiological substrates, biological roles and modes of
regulation of human HAD phosphatases, including the
functions of their numerous splice variants and disease-associated SNPs, will provide a rich field for further study.
Allosteric regulatory sites have begun to be identified in some HAD phosphatases. Together with more
structures of human HAD phosphatases becoming
available, this knowledge should greatly facilitate the
design of specific HAD phosphatase inhibitors for
potential future in vivo use.
FEBS Journal 280 (2013) 549–571 ª 2012 The Authors Journal compilation ª 2012 FEBS
563
564
MDP1
CTDP1
MDP-1
RNA polymerase
C-terminal
domain
phosphatases
EPHX2
NT5C1A
Soluble epoxide
hydrolase
cN-I nucleotidases
NT5C1B
PNKP
Polynucleotide
kinase
phosphatase
CTDNEP1
CTPSPL
CTDSP2
CTDSP1
Gene
HAD family
Cytosolic 5¢-nucleotidase
IA (cN-IA)
Cytosolic 5¢-nucleotidase
IB (cN-IB)
Soluble epoxide hydrolase
2 (sEH2)
Polynucleotide 5¢-kinase
3¢-phosphatase
Magnesium-dependent
phosphatase (MGDP-1);
fructosamine-6-phosphatase
(FN6Pase)
RNA polymerase II subunit
A C-terminal domain (CTD)
phosphatase;
TFIIF-associating CTD
phosphatase (Fcp1a)
Small C-terminal domain
phosphatase 1 (Scp1);
nuclear LIM interactor
interacting factor 3 (NLIIF)
small C-terminal domain
phosphatase 2 (Scp2);
conserved gene amplified
in osteosarcoma (OS4)
small C-terminal domain
phosphatase-like (Scp3)
CTP nuclear envelope
phosphatase; dullard
homolog
Proteins, synonyms
Table 1. Functionally characterized human HAD phosphatases.
2p24.2
1p34.3-p33
8p21.2-p21.1
19q13.4
17p13.1
3p22-p21.3
12q13-q15
2q35
18q23
14q12
Chrom.
localization
C1 cap, homotetramer
(NT5C1A)
C1 cap, homodimer
C0 cap, homodimer
PDB: 3ZVL (M. m.)
C0 cap
n.d.
C0 cap
PDB: 1T9Z
(H. s.)
PDB: 2Q5E
(H. s.)
C0 cap, monomer,
PDB: 1ONV (H. s.)
C0 cap, monomer,
PDB: 1U7O (M. m.)
Structure
AMP
Dihydroxy lipid
phosphates,
isoprenoid
phosphates
3¢-phosphorylated
DNA termini
pSer106-lipin1 [108]
Intracellular production
of adenosine under
ischemic and hypoxic
conditions
Lipid metabolism,
cholesterol
biosynthesis
Nuclear membrane
morphology; lipin1
subcellular targeting;
BMP signalling [109]
Multiple DNA repair
pathways
Regulation of gene
expression; BMP
and TGFb signaling;
cell migration
Regulation of
gene expression
pSer2 ⁄ pSer5 RNA
polymerase II
pSer5 RNA polymerase
II, pSmad, pSnail.
Scp3: pRB
Potentially involved in
protein glycation
repair [107]
Functions
Protein-bound
fructosamine-6phosphate
Substrates
Early infantile epileptic
encephalopathy-10
(OMIM #613402);
resistance to genotoxic
therapeutic agents
Dyslipidemia, coronary
heart disease, hypertension. Modifier of
familial hypercholesterolemia (OMIM
#143890) and of ischemic
stroke risk (OMIM
#601367)
Scp inhibition may
promote neuronal
regeneration; Scp2:
development of
sarcomas, Scp3:
tumor suppressor
Congenital cataracts
facial dysmorphism
neuropathy syndrome
(OMIM #604168)
Roles in disease
Human HAD phosphatases
A. Seifried et al.
FEBS Journal 280 (2013) 549–571 ª 2012 The Authors Journal compilation ª 2012 FEBS
NT5C2
cN-II nucleotidases
FEBS Journal 280 (2013) 549–571 ª 2012 The Authors Journal compilation ª 2012 FEBS
Eyes absent
Pyrimidine
5¢-nucleotidase ⁄
phosphoserine
phosphatases
Cytosolic 5¢-nucleotidase
II (cN-II); purine
5¢-nucleotidase; IMP-GMP
specific nucleotidase;
high Km 5¢-nucleotidase
Cytosolic 5¢-(3¢)deoxyribonucleotidase
(cdNT); uridine
5¢- monophosphate
hydrolase 2 (UMPH2)
Mitochondrial 5¢-(3¢)deoxyribonucleotidase
(mdNT)
Cytosolic 5¢nucleotidase III; uridine
5¢- monophosphate
hydrolase 1 (UMPH1);
pyrimidine 5¢nucleotidase (P5N-I)
Phosphoserine
phosphatase, PSP
Proteins, synonyms
PHOSPHO Phosphatase, orphan
1,2
1; phosphoethanolamine ⁄
phosphocholine (PEA ⁄
PCho) phosphatase isoform
1; phosphatase, orphan
2; pyridoxal phosphate
phosphatase PHOSPHO2
EYA1-4
Eyes absent 1 (Eya1)
Eyes absent 2 (Eya2)
Eyes absent 3 (Eya3)
Eyes absent 4 (Eya4)
PSPH
NT5C3
NT5M
NT5C
Deoxyribonucleotidases
Gene
HAD family
Table 1. (Continued)
Substrates
8q13.3
20q13.12
1p35.3
6q23.2
2q31.1
17q21.32
7p11.2
7p14.3
17p11.2
17q25.1
Organ development;
DNA-damage repair;
cell proliferation
Serine biosynthesis
Pyrimidine
catabolism in
reticulocytes
Regulation of
cellular dNTP pools
Regulation of cellular
dNTP pools
Purine nucleotide
interconversions and
IMP ⁄ GMP pools
Functions
Hemolytic anemia
(OMIM #266120)
Mediates resistance
against nucleoside
analogs used in cancer
and viral diseases
Roles in disease
EYA1: branchio-oto-(renal)
dysplasia syndromes
(OMIM #602588, #113650);
EYA4: dilated
cardiomyopathy-1J
(OMIM #605362), autosomal
dominant deafness-10
(OMIM #601316)
PSPH deficiency
syndrome (OMIM
#614023); ER-negative
breast cancer; PSPH
inhibition may promote
neuronal regeneration
PHOSPHO1: PEA,
PHOSPHO1 in chondrocytes: Potential therapeutic
PCho; PHOSPHO2:
generation of Pi for matrix
target in skeletal
pyridoxal 5¢-phosphate [110] mineralization
abnormalities with
skeletal development
excess or premature
mineral formation
L-3-phosphoserine
5¢- dUMP, 5¢-, 3¢dTMP, 5¢-, 3¢-,
2¢- UMP
UMP, CMP
All 5¢-dNMPs
except dCMP
C1 cap
Eya3: pTyr-142-H2A.X
PDB: ED-EYA2 3GEB;
3HB0; 3HB1
(H. s.)
C1 cap, additional
small C2 cap
C1 cap, homodimer
PDB: 1L8L (H. s.)
C1 cap, homodimer
PDB: 2JAW
(H. s.)
C1 cap, monomer
PDB: 2VKQ (H. s.)
C1 cap, homodimer
PDB: 2JAR
(M. m.)
10q24.32- C1 cap, homotetramer IMP, GMP
q24.33
PDB: 2J2C
(H. s.)
Chrom.
localization Structure
A. Seifried et al.
Human HAD phosphatases
565
566
PMM1, -2
LPIN1-3
Cof hydrolases
Lipins
PDXP
Lipin 1-3,
phosphatidase phosphatase
(PAP), phosphatidic acid
hydrolase (PAH)
lipin1
lipin2
lipin3
Phosphomannomutase1
phosphomannomutase2
Phospholysine
phosphohistidine inorganic
pyrophosphate phosphatase
LHPP
NagD
NANP
Haloacid dehalogenase-like
hydrolase domain containing
1A; pseudouridine-5¢monophosphatase
(5¢-PsiMPase)
N-acylneuraminate-9phosphatase; haloacid
dehalogenase-like
hydrolase domain-containing
protein 4 (HDHD4)
Pyridoxal 5¢-phosphate (PLP)
phosphatase, Pdxp,
chronophin (CIN)
HDHD1
HAD-like
hydrolase
domaincontaining
proteins
Proteins, synonyms
Gene
HAD family
Table 1. (Continued)
2p25.1
18p11.31
20q12
22q13.2
16p13.2
10q26.13
22q12.3
20p11.2p11.1
Xp22.31
Chrom.
localization
C2 cap, homodimer,
PDB: 2FUE (PMM1);
2AMY(PMM2) (H. s.)
Homodimer or -tetramer;
hetero-oligomers
C2 cap,
homodimer
PDB: 2OYC; 2P69
(H. s.)
C2 cap, homodimer
PDB: 2X4D (H. s.)
C1 cap
PDB: 2W4M
(H. s.)
C1 cap
PDB: 3L5K
(H. s.)
Structure
PMM1: Glc-1,6-P2
(IMP-stimulated); PMM2:
mannose-6-phosphate
Phosphatidic acid
PLP [113]; pSer3-cofilin;
steroid receptor
coactivator 3
(pSer ⁄ pThr) [114]
N-acetyl-neuraminate9-phosphate [112]
Pseudouridine 5¢phosphate [111]
Substrates
PMM1: ischemic brain
metabolism; PMM2:
protein N-glycosylation
Lipid metabolism
Vitamin B6 metabolism;
regulation of actin
cytoskeletal dynamics;
estrogen receptor signaling
Sialic acid formation: roles
in protein-protein and
cell-cell recognition
Pseudouridine excretion
upon RNA breakdown
Functions
HTR1A and LHPP SNPs
are interacting genetic
risk factors in major
depression [115]
PMM2: congenital
disorder of glycosylation,
type Ia (OMIM #212065)
LPIN1: rhabdomyolysis
(OMIM #268200);
statin-induced myopathy;
LPIN2: Majeed syndrome
(OMIM #609628),
psoriasis
Potentially involved
in breast cancer
Often deleted in X-linked
ichthyosis (OMIM
#300747)
Roles in disease
Human HAD phosphatases
A. Seifried et al.
FEBS Journal 280 (2013) 549–571 ª 2012 The Authors Journal compilation ª 2012 FEBS
A. Seifried et al.
Human HAD phosphatases
Acknowledgements
This work was supported by grants from the DFG
(SFB688, to A.G.), and by the Rudolf Virchow Center
(DFG ⁄ FZ82, to A.G.).
13
14
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Supporting information
The following supplementary information is available:
Table S1. Protein-coding transcripts of human HAD
phosphatases.
Doc. S1. Methods to determine the evolutionary history of metazoan HAD phosphatases.
This supplementary material can be found in the
online version of this article.
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